Constitutive expression of cytochrome P450 1B1 endows testicular Leydig cells with susceptibility to 7,12-dimethylbenzanthracene-induced cell death

Abstract

Testicular Leydig cells produce testosterone through the participation of steroidogenic proteins. The CYP1B1 enzyme has been shown to catalyze 7,12-dimethylbenzanthracene (DMBA), a representative polycyclic aromatic hydrocarbon. We hypothesized that exposure to DMBA causes Leydig cell cytotoxicity through activation of CYP1B1. Leydig cells were exposed to various concentrations of DMBA for the induction of CYP1B1 expression and activity. The status of CYP1B1 function was monitored by evaluation of cytotoxicity-mediated cell death. Our data show that exposure to DMBA causes cytotoxicity in Leydig cells by CYP1B1 activation. DMBA evoked a significant increase in the generation of reactive oxygen species (ROS) by which the depolarization of mitochondrial membrane potential (MMP) is initiated and caspase-3 activation is augmented. The knockdown of CYP1B1 expression resulted in the suppression of DMBA-induced apoptosis via reduced p53 activation and caspase-3 activation, suggesting that a final metabolite of DMBA (i.e., DMBA-DE) bioactivated by CYP1B1 induces p53 activation by binding to DNA and subsequently causing apoptosis via caspase-3 activation. This finding provides evidence for constitutive expression of CYP1B1 in Leydig cells, which is a trait that only requires an initiating signal for its activity. Further research on CYP1B1 activation-provoked steroid metabolism in Leydig cells may provide decisive clues for elucidating its innate function.

INTRODUCTION

Leydig cells, which are testosterone-producing cells, are located in the interstitial compartment of the testes. Testosterone plays a crucial role in the maintenance of spermatogenesis (Roberts and Zirkin, 1991). Testosterone is synthesized in Leydig cells through the participation of important steroidogenic proteins, such as steroidogenic acute regulatory protein (StAR), cytochrome P450 side-chain cleavage (P450scc), and 3β-hydroxysteroid dehydrogenase isomerase (3β-HSD). Aberrant changes in Leydig cell functions can provoke abnormalities in male reproduction processes, primarily by disruption of testosterone production. Exposure to various environmental byproducts and/or food contaminants can cause testicular toxicity by disturbing Leydig cells endocrine functions (Akingbemi et al., 2004; Culty et al., 2008; Hancock et al., 2009; Chung et al., 2011). Toxicant-evoked endocrine disruptions in Leydig cells appear to primarily occur as a result of steroidogenic protein dysregulation (Culty et al., 2008; Chung et al., 2011). However, other potential mechanism(s) associated with cytotoxicity-mediated Leydig cell dysfunction must be further elucidated.

Among cellular enzymes, the cytochrome P450 (CYP) superfamily of monooxygenases is a large and diverse group of enzymes that metabolize organic substances, including lipids, steroid hormones, xenobiotic drugs, and other toxic chemicals (Omura, 1999). To date, 57 active CYP genes have been identified in the human genome and are divided into 18 families (Nebert and Dalton, 2006). Depending on cellular conditions, CYPs will play a major role in either the detoxification or the bioactivation of toxic substances. Thus, if a specific xenobiotic toxicant is metabolized toward bioactivation within a cell, it could damage cellular components and subsequently alter the cells’ fate toward either carcinogenesis or apoptosis. The potential role of several CYP enzymes in Leydig cells has been studied (Simpson et al., 1994; Payne and Youngblood, 1995; Chung et al., 2007). Although CYP enzymes associated with steroidogenesis, including CYP11A1, CYP17A1, and CYP19, are well recognized in Leydig cells, the potential involvement of other CYP enzymes is poorly understood. Among the enzymes that are not well understood, greater attention should be paid to understanding the role of CYP1B1 because this enzyme is constitutively expressed in Leydig cells (Otto et al., 1992; Walker et al., 1995). While some effort has been made to study the potential roles of CYP1B1 in testicular Leydig cells (Leung et al., 2009; Deb et al., 2010), a genuine role for CYP1B1 in Leydig cells still remains unknown.

CYP1B1 is abundantly expressed in the kidney, prostate, breast, and uterus of humans (Shimada et al., 1996). The CYP1B1 enzyme has been shown to catalyze the regioselective modification of 7,12-dimethylbenzanthracene (DMBA) (Pottenger and Jefcoate, 1990; Otto et al., 1992), a representative polycyclic aromatic hydrocarbon (PAH), that exhibits carcinogenic, teratogenic, and apoptogenic properties (Cavalieri et al., 1991; Chidambaram and Baradarajan, 1996; Yamaguchi et al., 1997). Additionally, DMBA has been identified as a selective inducing factor for the upregulation of CYP1B1 expression and activity in human endometrial cancer cells (Kim et al., 2012). DMBA is therefore an ideal agent for studying CYP1B1 function, in addition to being an important PAH whose cytotoxicity should be assessed in cells with constitutively expressed CYP1B1.

Here, we hypothesized that exposure to DMBA causes Leydig cell cytotoxicity through activation of CYP1B1. Leydig cells were exposed to DMBA for the induction of CYP1B1 expression and activity. The status of CYP1B1 function was monitored by evaluation of cytotoxicity-mediated cell death.

MATERIALS AND METHODS

Reagents and antibodies

7,12-Dimethylbenzanthracene (DMBA), dimethyl sulfoxide (DMSO), 3-[4,5-dimethylthiazol-2-yl]-2.5-diphenyltetrazolium bromide; thiazolyl blue (MTT), propidium iodide (PI), Rhodamine 123, α-naphthoflavone, pyrene, pifithrin-α, 5,11-dimethyl-6H-pyrido[4,3-b]carbazole (ellipticine), phenylmethyl-sulfonyl fluoride (PMSF), CYP1B1 isozyme, and anti-actin antibody were purchased from Sigma-Aldrich (St. Louis, MO, USA). 2,3′,4,5′-tetramethoxystilbene (TMS) was from Cayman Chemical (Ann Arbor, MI, USA). Dichlorofluorescein diacetate (DCF-DA) fluorescent probe was from Invitrogen Corporation (Carlsbad, CA, USA). RNase A was from Quigen (Valencia, CA, USA). Anti-cytochrome c, p53, CYP1A1, and CYP1B1 antibodies, and mouse CYP1B1 siRNA and Control siRNA-A were obtained from Santa Cruz Biotech (Santa Cruz, CA, USA). Anti-cleaved caspase-3, phospho-p-53(ser-15) antibodies, rabbit and mouse IgG-conjugated with horseradish peroxidase were purchased from Cell Signaling (Beverly, MA, USA). The pancaspase inhibitor (zVAD-fmk) and capsase-3 inhibitor (DEVD-CHO) were from Calbiochem (San Diego, CA, USA). FITC-conjugated anti-rabbit IgG and anti-mouse IgG were from Vector Laboratory (Burlingame, CA, USA).

Leydig cell culture

TM3 (normal mouse Leydig cell line; American Type Culture Collection, Manassas, VA, USA) were cultured in Dulbecco’s modified Eagle’s medium nutrient mixture F-12 HAM (Sigma-Aldrich) containing with 5% heat-inactivated horse serum (Invitrogen), 2.5% heat-inactivated fetal bovine serum (Invitrogen) and 1.2 g/L sodium bicarbonate supplemented with 10 µg/mL penicillin-streptomycin (Invitrogen). The cells (5~15 passages) were incubated at 37ºC in a humidified incubator with an atmosphere of 5% CO₂ and were exposed to DMBA when confluency reached 30%.

Knockdown of CYP1B1 by small interfering RNA (siRNA) transfection

Cells were transfected with mouse CYP1B1 siRNA (Santa Cruz) by using siRNA transfection reagent (Santa Cruz) according to the manufacturer’s suggested protocol. Briefly, a defined number of cells (4 × 10⁵) were incubated in 6-well plate for 18~24 hr before transfection. The cells were monolayerly 50~60% confluent at the time of transfection. Cells were washed once with serum and antibiotics-free medium [or siRNA transfection medium (Santa Cruz)] and subsequently transfected 40 pM of either mouse CYP1B1 siRNA (Santa Cruz, sc-44547) or control siRNA (Santa Cruz, sc-37007) for 8 hr at 37ºC. After transfection, the cells were cultivated in normal growth medium without removing the transfection mixture and exposed to DMBA. At 24, 48 and 72 hr after DMBA treatment, cells were harvested and then were assessed by cell death and protein analysis.

MTT cell viability assay

Cells were seeded in 12-well plates at a density of 5 × 10⁵ cells per well. After treatment at an appropriate time, the culture medium was removed and replaced with a medium containing 0.5 mg of MTT dissolved in PBS (pH 7.2), After 4 hr, the formed crystals were dissolved with 200 µL of DMSO. The intensity of the color in each well was measured at a wavelength of 490 nm using a microplate reader (BIOTEK EL-312e, VT, USA).

Flowcytometric cell death assay

The cells were harvested, fixed with 95% ethanol for 24 hr, incubated with 0.05 mg/mL PI and 1 μg/mL RNase A at 37ºC for 30 min, and analyzed by flow cytometry, using an Epics XL and analysis software (EXPO32^TM; Beckman Coulter, MI, USA). The cells belonging to the sub-G1 population were considered to be apoptotic cells; the percentage of each phase of the cell cycle was determined.

Western blot analysis

Whole-cell lysates were prepared by incubating cell pellets in lysis buffer [30 mM NaCl, 0.5% Triton X-100, 50 mM Tris-HCl (pH 7.4), 1 mM Na₃VO₄, 25 mM NaF, 10 mM Na₄P₂O₇] for 30 min on ice. After the insoluble fractions were removed by centrifugation at 20,800 × g at 4ºC for 30 min, the supernatants were collected and protein concentration was determined with a BCA protein assay kit (Pierce Biotechnology, Woburn, MA, USA). The same amounts of proteins (~30 μg) in SDS-gel sample buffer [2% (w/v) SDS, 100 mM DTT, 10% (v/v) glycerol, 0.01% (w/v) bromophenol blue, and 60 mM Tris-HCl, pH 6.8] were boiled at 95ºC for 10 min and resolved by 7–15% gradient SDS-PAGE. For Western blots, the gels were subsequently equilibrated in transfer buffer [26 mM Tris, 192 mM glycine, 20% (v/v) methanol, pH 8.3], and the separated proteins were transferred to a 0.2-μm nitrocellulose membrane in a Tris/glycine transfer buffer containing 10% methanol by using a Mini Trans-Blot Cell Western blotting apparatus (Bio-Rad, Hercules, CA, USA) at 100 V for 1 hr at 4ºC. Nonspecific binding sites were blocked with 5% (w/v) nonfat dry milk in TBS-T (25 mM Tris, 137 mM NaCl, 3 mM KCl, and 0.05% Tween-20). The blots were incubated for 1 hr each at 37ºC in primary and appropriate HRP-conjugated secondary antibodies diluted in TBS-T according to the manufacturer’s instructions. The membrane was then washed in TBS-T, followed by washing in TBS alone. The signals were detected with an enhanced chemiluminescence detection kit (Amersham Pharmacia Biotech, Piscataway, NJ, USA) in the LAS-3000 detector (Fujifilm, Japan). Immunoblotting for β-actin was performed in every experiment as an internal control.

Immunocytochemistry

Harvested cells were attached on the slide glass by cytospin centrifugation. The cells were fixed with 4% PFA, washed with PBS, and incubated with 0.2% Triton X-100. Then, the cells were incubated with the appropriate primary antibody in 1% bovine serum albumin at room temperature (RT). For secondary antibody reaction, the cells were incubated with an appropriate fluorescence-conjugated secondary antibody at RT. For counterstaining of the nucleus, if required, cells were incubated with PI (50 μg/mL) at RT. Finally, cells were mounted and observed under a confocal microscope (LSM700, Carl Zeiss, Germany) at the Neuroscience Translational Research Solution Center (Busan, South Korea).

Determination of intracellular ROS production

Reactive oxygen species (ROS) generation was monitored by staining cells with DCF-DA. Following exposure to DMBA, cells were incubated with 5 μM DCF-DA at 37ºC for 30 min. The cells were trypsinized, washed with PBS, suspended in PBS, and analyzed using flow cytometry.

Measurement of mitochondrial membrane potential (MMP)

The cells (5 × 10⁵) were incubated with 5 μM Rhodamine 123 and 50 nM Mitotraker CMXRos dye at 37ºC for 30 min, washed and resuspended with PBS, and then the fluorescence [red (585/590 nm); green (510/527 nm)] was measured by flow cytometer.

Caspase-3 activity assay

A fluorometric assay kit (Clontech, CA, USA), which contains fluorogenic substrates specific for caspase-3 immobilized in the wells, was used to evaluate enzyme activity. Ten micrograms of the extracted proteins in homogenization buffer (50 mM Tris HCl, 150 mM NaCl, 10% glycerin, and 1% Triton X-100) were added to the wells. The plate was incubated in the fluorescence plate reader at 37ºC for 3 hr, and fluorescence was read every 10 min. The activity was determined by fluorometric detection (excitation, 380 nm; emission, 460 nm) and the negative control (blank, without sample) was subtracted from all the samples. Results at 2 hr were selected, as the manufacturer suggested. Baseline values of negative controls and samples with specific inhibitors did not increase during the 2-hr interval.

CYP1B1 enzyme activity assay

Cellular proteins were isolated with lysis buffer consisting of 30 mM NaCl, 50 mM Tris–HCl (pH 7.6), 5% Triton X-100, and 100 mM phenylmethylsulfsonyl (PMSF). The enzymatic activity of CYP1B1 was measured by P450-Glo™ assay kits (Promega; Madison, WI, USA) as per manufacturer’s instruction manual. Briefly, isolated proteins (30 μg) were mixed with the 4 × cytochrome P450/KPO₄/substrate reaction mixture (CYP1B1 buffer: 1 pM CYP1B1 isozyme, 400 mM KPO₄, 80 μM luciferin-CEE), and 2 × NADPH regeneration system (Promega). D-Luciferin was used to generate a standard curve, and the corresponding CYP1B1 isozyme was omitted from blank control. If required, a control (known) inhibitor was added to the reaction. The sample and the 4 × cytochrome P450/KPO₄/substrate reaction mixture were added to a 96-well plate. After preincubating the plate at 37ºC for 20 min, the 2 × NADPH regeneration system was added to each reaction. The plate was incubated at 37ºC for 30 min, and the reconstituted leuciferin detection reagent was added. Again, the plate at RT was incubated for 20 min, and the luminescence recorded using a Microplate Luminometer (GloMax-96; Promega).

Statistics

Data were expressed as the mean ± SD of 3 or 4 separate experiments. Where appropriate, the data were subjected to analysis of variance (ANOVA) followed by Tukey’s test. The means were considered significantly different at p < 0.05.

RESULTS

DMBA induces cell death in TM3 Leydig cells

Sub-G1 cell cycle analysis by flow cytometry showed that 10 μM DMBA induced significant cell death at 48 hr after treatment (Fig. 1A), and that prolonged treatment with DMBA resulted in increased cell death (Fig. 1B). A DMBA concentration of 10 μM was thus considered optimal for the time-course experiments with TM3 cells.

Fig. 1

Effects of 7,12-dimethylbenzanthracene (DMBA) on cell death in TM3 Leydig cells. A, Cell death shown in cells treated with various concentrations of DMBA (0.01~100 μM) for 48 hr and assessed by flow cytometry. B, Cell death shown in cells treated with 10 μM DMBA at different time-points, and assessed by flow cytometry. At least three independent experiments were performed and data shown are the mean ± SD. #, p < 0.001 compared with 0 μM or 0 hr control.

DMBA-induced cell death is accompanied by the generation of ROS, mitochondrial alterations, and caspase-3 activation

An increase in intracellular ROS is often linked to the occurrence of apoptotic cell death (Fleury et al., 2002). Exposure to DMBA provoked the generation of a significant amount of ROS at 48 and 72 hr, but not at earlier time points (12 and 24 hr) (Fig. 2A). Apoptosis is accelerated by a decrease in mitochondrial membrane potential (MMP) and the release of cytochrome c from the mitochondrial membrane into the cytosol (van Loo et al., 2002). Exposure to DMBA caused depolarization of MMP (Fig. 2B) and the release of cytochrome c into the cytosol (Fig. 2C). To determine whether DMBA-induced TM3 cell apoptosis occurs in a caspase-dependent manner, activation of caspase-3 was analyzed. Exposure to DMBA induced caspase-3 activation (Fig. 2D) and activated caspase-3 was markedly localized to DMBA-treated cells (Fig. 2E). DMBA-induced caspase-3 activity was suppressed in the presence of the pancaspase inhibitor zVAD-fmk and in the presence of the caspase-3 specific inhibitor DEVD-CHO (Fig. 2F).

Fig. 2

Changes in generation of intracellular reactive oxygen species (ROS), mitochondrial membrane potential (MMP, ∆Ψ_m), release of mitochondrial cytochrome c, and caspase-3 activation in TM3 Leydig cells following exposure to DMBA. A, Generation of ROS in TM3 cells following exposure to DMBA. Cells were treated with 10 μM DMBA for the indicated times and stained with DCF-DA for flow cytometric analysis. Thin line, control cells; short-dotted line, DMBA-treated cells. B, Flow cytograms of MMP (∆Ψ_m) in TM3 cells after exposure to DMBA. The cells were treated with DMBA (10 μM) for the indicated times, stained with Rhodamine 123, and measured by flow cytometry. Thin line, control cells; short-dotted line, DMBA-treated cells. Arrows indicate the depolarization status of MMP. C, Immunocytochemical localization of cytochrome c after exposure to DMBA. The cells were treated for 72 hr with DMBA (10 μM), cytospun, fixed, and immunostained with a cytochrome c antibody. Con and DMBA indicate control and 7,12-dimethylbenzanthracene, respectively. Microphotographs were taken using confocal microscopy. Original magnification: × 800. D, Western blot analysis of cleaved caspase-3 after exposure to DMBA. The cells were exposed to DMBA (10 μM) for 12, 24, 48, and 72 hr. Actin is indicated as an internal loading control. E, Immunocytochemical localization of active caspase-3 after exposure to DMBA. The cells were treated for 72 hr with DMBA (10 μM), cytospun, fixed, and immunostained with a active form-specific caspase-3 antibody. CON and DMBA indicate control and 7,12-dimethylbenzanthracene, respectively. Microphotographs were taken using confocal microscopy. Original magnification: × 800. F, Caspase-3 activity measured spectrofluorometrically using a caspase-3 specific substrate (Ac-DEVD-AMC), as described in Materials and Methods. TM3 cells were treated with 10 μM DMBA for 48 hr in the absence or presence of the pancaspase inhibitor (zVAD-fmk, 10 μM) or the caspase-3 specific inhibitor (Ac-DEVD-CHO, 0.5 μM). Con, DMBA, VAD+D, and DEVD+D indicate control, 7,12-dimethylbenzanthracene, zVAD-fmk + 7,12-dimethylbenzanthracene, DEVD-CHO + 7,12-dimethylbenzanthracene, respectively. At least three independent experiments were performed and data shown are the mean ± SD. +, p < 0.001 compared to control. * and ** indicate p < 0.05 and p < 0.001 compared to the DMBA-treated group, respectively.

Constitutive levels of CYP1B1 expression in Leydig cells are enhanced by exposure to DMBA

Changes in the levels of CYP1B1 and CYP1A1 proteins were monitored to determine whether CYP1B1 expression is significantly altered following exposure to DMBA. While CYP1B1 protein levels in the control cells remained considerably high, CYP1A1 expression remained at basal levels throughout the culture period (Fig. 3A). DMBA exposure resulted in increased CYP1B1 protein levels in TM3 cells during treatment (Fig. 3A and 3B). CYP1A1 protein levels also appeared to increase in response to DMBA treatment; however, the CYP1A1 protein increase was negligible when compared with CYP1B1 protein (Fig. 3A and 3B).

Fig. 3

Changes in CYP1B1 and CYP1A1 expression for TM3 Leydig cells following exposure to DMBA. A, Western blot analyses of CYP1B1 and CYP1A1 protein expression. The cells were treated at different time points with DMBA (10 μM). Actin is indicated as an internal loading control. CON and DMBA indicate control and 7,12-dimethylbenzanthracene, respectively. B, Immunolocalization of CYP1B1 and CYP1A1. Cells were treated for 48 hr with 10 μM DMBA, cytospun, fixed, and immunostained with antibodies as indicated. The images were taken by a confocal microscope. Original magnification: × 800.

CYP1B1 activation is required for DMBA-induced apoptosis in TM3 Leydig cells

CYP1B1 activity and cell death were measured in DMBA-treated TM3 cells in the absence or presence of various CYP1 inhibitors to investigate whether DMBA-induced upregulation of CYP1B1 expression is associated with CYP1B1 activity, as well as with cytotoxicity-mediated apoptotic cell death. In this study, we consistently observed that CYP1B1 activity in Leydig cells remained endogenously elevated (Fig. 4A). Although DMBA treatment increased CYP1B1 activity (~1.5 fold) compared to that for the control, the increased ratio was in fact lower than our predicted level (Fig. 4A). The CYP1B1-specific activity assay showed that CYP1B1 activity was significantly reduced by α-NF (a pan-CYP1 inhibitor); activity was also suppressed by pyrene and pifithirin-α (PFT-α; CYP1B1-specific inhibitors) but not by the CYP1A1-specific inhibitor ellipticine (Fig. 4A). However, among the CYP1B1 inhibitors, TMS failed to suppress CYP1B1 activity (Fig. 4A). We found that ellipticine and TMS are not capable of suppressing DMBA-induced apoptosis (Fig. 4B and 4C). The results displayed in Figure 4C further show that pretreatment with α-NF, pyrene, and PFT-α resulted in the suppression of DMBA-induced apoptosis by preventing p53 phosphorylation and caspase-3 activation.

Fig. 4

Suppression of CYP1B1 activity and cell death in TM3 Leydig cells following exposure to DMBA. In A, B, and C, cells were either treated with DMBA (10 μM) or vehicle (0.1% DMSO) for 72 hr in the absence or in the presence of α-NF (ANF, α-naphthoflavone, 1 μM), TMS (2,3′,4,5′-tetramethoxystilbene, 0.1 μM), pyrene (PYR, 50 μM), PFT-α (PFT, pifithrin-α, 10 μM), or Ellipiticin (ELL, 1 μM). At least three independent experiments were performed and data shown are the mean ± SD. +, p < 0.001 compared to control. *, **, and # indicate p < 0.05, p < 0.01, and p < 0.001 compared to the DMBA-treated group, respectively. A, Change in DMBA-induced CYP1B1 activity in the absence or presence of CYP1 inhibitors, measured by fluorogenic specific enzyme activity assays, as described in Materials and Methods. B, Cell death determined by sub-G1 analysis of flow cytometry. C, Changes in CYP1B1, CYP1A1, p53, phospho-p53, and cleaved caspase-3 protein detected by Western blotting. Actin is indicated as an internal loading control. In D, E, and F, for knockdown of CYP1B1 gene in TM3 cells, si-RNA for CYP1B1 or control siRNA was transfected for 8 hr prior to exposure to DMBA (10 μM) for 72 hr. *, **, and # indicate p < 0.05, p < 0.01, and p < 0.001 compared to each indicated group, respectively. D, Changes in CYP1B1 activity resulting from CYP1B1 knockdown in the absence or presence of DMBA. CYP1B1 activity was measured by fluorogenic enzyme-specific activity assays, as described in Materials and Methods. E, Cell death determined by sub-G1 analysis of flow cytometry. F, Changes in CYP1B1, CYP1A1, p53, phospho-p53, and cleaved caspase-3 protein by CYP1B1 knockdown, as shown by Western blots. Actin is indicated as an internal loading control.

Knockdown of CYP1B expression blocks DMBA-induced apoptosis in TM3 Leydig cells

Based on the aforementioned data, we postulated that CYP1B1 is the key CYP1 enzyme for DMBA-induced cytotoxicity during TM3 cell apoptosis. To examine this hypothesis, we employed an anti-CYP1B1 si-RNA system. Constitutively expressed CYP1B1 levels in TM3 cells did not appear to be suppressed by introduction of the CYP1B1 si-RNA (Fig. 4D). However, the elevated CYP1B1 activity induced by DMBA was significantly suppressed by CYP1B1 knockdown (Fig. 4D). Furthermore, DMBA-induced apoptosis was markedly suppressed in CYP1B1-knockdown cells (Fig. 4E). CYP1B1 knockdown also reduced the p53 phosphorylation and caspase-3 activation that are typically induced by DMBA (Fig. 4F).

DISCUSSION

PAHs are toxic environmental contaminants that are produced during incomplete combustion of organic materials. The major environmental sources of human exposure include cigarette smoke, barbecued food, and automobile exhaust. Among PAHs, DMBA is one of the most potent mutagenic and carcinogenic compounds, as it has been associated with the induction of tumors in breast tissue and skin (Cavalieri et al., 1991; Chidambaram and Baradarajan, 1996). The apoptogenic capability of DMBA has been demonstrated in bone marrow B cells (Heidel et al., 1999), ovarian cells (Matikainen et al., 2001), ovarian preovulatory follicles (Tsai-Turton et al., 2007) and uterine endometrial cells (Kim et al., 2012). However, the cytotoxicity and apoptosis provoked by DMBA has not yet been investigated in testicular Leydig cells.

We have previously shown that exposure to benzo[a]pyrene (B[a]P), another representative PAH, does not cause cell death in TM3 Leydig cells (Chung et al., 2007), because B[a]P is not able to evoke cell death by conversion to cytotoxic (DNA adductable) structures (i.e., B[a]P-7,8-dihydroxy-9-10-epoxide [BPDE]) in the case of CYP1A1 insufficiency in TM3 cells. In contrast, in the present study, TM3 Leydig cells underwent cell death after exposure to DMBA. These outcomes indicate that DMBA can exhibit its cytotoxicity in Leydig cells either directly by itself, or through bioactivation without involvement of CYP1A1. Following exposure to B[a]P or DMBA, several cell types exhibit caspase-dependent apoptosis accompanied by mitochondrial alterations (Matikainen et al., 2002; Kim et al., 2007; Chung et al., 2007; Kim et al., 2012). Consistent with previous findings, we showed that DMBA-induced apoptosis in TM3 Leydig cells is accompanied by activation of caspase-3. In addition, mitochondrial changes ―the decrease in membrane potential and release of cytochrome c into the cytosol―indicate that the mitochondrial pathway is closely associated with DMBA-induced apoptosis. Accumulation of ROS is often linked to the occurrence of apoptotic cell death (Fleury et al., 2002). In this study, DMBA evoked a significant increase in the generation of ROS by which the depolarization of MMP is initiated and caspase-3 activation is augmented. Thus, ROS are likely not a prompt initiator of cell death in DMBA-treated Leydig cells, but may contribute to the enhanced apoptosis provoked by MMP alteration.

For initiation of cytotoxicity-induced apoptosis by PAHs, these chemicals must be modified to forms that can readily bind DNA or provoke DNA damage. The CYP1 family of enzymes (CYP1A1, CYP1A2, and CYP1B1) are responsible for these processes with PAHs (Alexander et al., 1997; Kleiner et al., 2004); therefore, upregulation of CYP1 expression and activity is required. DMBA is biotransformed in cells into its cytotoxic form, DMBA-DE, by 3 sequential reactions mediated by CYPs and mEH (Shimada and Fujii-Kuriyama, 2004). CYP1B1 predominantly catalyzes epoxidation at the 3,4-positions of DMBA. mEH then hydrolyzes DMBA-3,4-epoxide to DMBA-3,4-diol, which is then oxidized again, by either CYP1A1 or CYP1B1, to the principal carcinogen DMBA-DE. This covalently binds to DNA, forming a DMBA-DE-DNA adduct (Cheng et al., 1988), and causes DNA damage, thereby causing cytotoxicity. Therefore, CYP1B1 activation is considered essential for eliciting DMBA activity in cellular systems (Kim et al., 2012).

In the present study, we have shown that CYP1B1 is significantly upregulated in TM3 Leydig cells compared with control cells following exposure to DMBA. In contrast, a study showed that CYP1B1 expression is not inducible in Leydig cells by B[a]P (Deb et al., 2010). This finding implies that CYP1B1 in Leydig cells is expressed in a ligand-selective manner. CYP1A1 appears to be slightly increased by exposure to DMBA in TM3 cells. However, considering the fact that normal Leydig cells isolated from the testis are deficient for CYP1A1 gene expression (Chung et al., 2007), the involvement of CYP1A1 in DMBA metabolism in Leydig cells could indeed be excluded. An important observation resulting from this study was that the basal level of CYP1B1 activity in Leydig cells remains noticeably high. Thus, although exposure to DMBA significantly increased CYP1B1 activity, the extent of increase was consistently less than ~1.5 fold when compared with that for the control. Nevertheless, the rate of cell death provoked by the increased activity was significant. This finding may provide evidence for constitutive expression of CYP1B1 in Leydig cells, which is a trait that only requires an initiating signal for its activity.

The association of CYP1B1 activity in DMBA-induced apoptosis was supported by the CYP1 inhibitor-screening test that was performed in this study. α-NF was introduced into cell cultures as a pan-CYP1 inhibitor (Shimada et al., 1998; Kim et al., 2012); TMS (Chun et al., 2001), pyrene (Shimada et al., 1998), and PFT-α (Sparfel et al., 2006), as CYP1B1-selective inhibitors; and ellipticine (Tassaneeyakul et al., 1993), as a selective CYP1A1 inhibitor. α-NF, pyrene, and PFT-α suppressed DMBA-induced CYP1B1 activity, but ellipticine failed to inhibit the enzyme activity. This observation indicates that CYP1B1 activity enhanced by DMBA is comparatively specific. However, among the specific CYP1B1 inhibitors, the TMS failed to suppress CYP1B1 activity as well as cell death induced by DMBA. The nature of this phenomenon is currently unknown; however, it is possible that the inhibitory action of TMS could be either tissue (or cell) type-specific or species-specific. The apparent involvement of CYP1B1 activity in DMBA-induced apoptosis in Leydig cells was further confirmed by the CYP1B1 gene knockdown experiment. The knockdown of CYP1B1 expression resulted in the suppression of DMBA-induced apoptosis via reduced p53 activation (phosphorylation) and caspase-3 activation, suggesting that a final metabolite of DMBA (i.e., DMBA-DE) bioactivated by CYP1B1 induces p53 activation by binding to DNA and subsequently causing apoptosis via caspase-3 activation. Therefore, p53 is a prerequisite for the induction of apoptosis in cells that are affected by DMBA.

Our data show that exposure to DMBA causes cytotoxicity in Leydig cells by CYP1B1 activation. Although the precise role of CYP1B1 in Leydig cells remains currently unknown, further research on CYP1B1 activation-provoked endocrine changes and steroid metabolism in Leydig cells may provide decisive clues for elucidating its innate function.

ACKNOWLEDGMENTS

This work was supported by Korea Environment Industry & Technology Institute (KEITI) through Technology Development Project for Safety Management of Household Chemical Products, funded by Korea Ministry of Environment (MOE) (1485017593).

Conflict of interest

The authors declare that there is no conflict of interest.

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